Translational control allows cells to rapidly change their proteome to respond to external triggers or other cues without de novo mRNA synthesis (10, 11). In fact, modulation of translational efficiency represents a critical mechanism in a plethora of biological processes, such as cell differentiation, metabolism, growth, and proliferation (10, 12–14). Accordingly, dysregulation of mRNA translation is a hallmark of various types of cancer (15, 16) and other clinical disorders, such as inflammatory airway pathologies (17), fibrosis (18), and neurodegenerative diseases (19–22). In eukaryotes, translation is mainly regulated at the initiation step, during which ribosomes are recruited to the mRNA, a process which can be modulated via multiple mechanisms. For instance, the association of mRNAs with RNA-binding proteins (23) and the presence of features including the 5′-terminal oligopyrimidine (5′ TOP) motif (24) or structured sequence motifs within the 5′ untranslated region (UTR) of mRNA (25) represent regulatory mechanisms selectively influencing translational efficiency. Notably, ribosome recruitment is facilitated by the recognition of the mRNA 5′-m7G cap structure by eukaryotic initiation factor 4E (eIF4E), which, together with the scaffold protein eIF4G and the RNA helicase eIF4A, forms the eIF4F complex (26). Assembly of the eIF4F complex is prevented by eIF4E-binding proteins (4E-BPs), which block the eIF4E-eIF4G interaction and eIF4F formation (27, 28). Hyperphosphorylation of 4E-BPs by the serine/threonine kinase mechanistic target of rapamycin (mTOR) complex 1 (mTORC1) leads to a reduction of the 4E-BPs' affinity for eIF4E, which favors the eIF4E-eIF4G interaction and the initiation of translation (29). Thus, signaling through the mTORC1 axis is pivotal for translational control.

Regulation of mRNA translation efficiency is required for normal immune functions (30) and is altered during infection (31, 32). mTORC1 and its downstream targets 4E-BP1/4E-BP2 and S6K1/S6K2 are key components of the innate immune response (33–37). Accordingly, changes in mTORC1 signaling are linked to subversion of host mRNA translation by viruses (38, 39), bacteria (40, 41), and the protozoan parasite Leishmania major (42).

With regard to toxoplasmosis, translational control has been assessed only in the parasite (43, 44); however, how the parasite modulates host cell translation is unknown. Here, we report that T. gondii selectively reprograms the translational landscape of the host cell to promote its replication. Through a nonbiased approach (i.e., transcriptome-wide polysome profiling), we identified a large number of transcripts whose translation efficiency is modulated upon infection (i.e., activated or repressed). In addition, we show that selective activation of host mRNA translation by T. gondii is mTOR sensitive and is associated with the presence of distinct motifs in the 5′ UTRs of identified transcripts. Accordingly, inhibition of mTOR activity dampens parasite replication. Overall, this study provides evidence that selective regulation of host mRNA translation contributes to T. gondii survival.

RESULTS

Toxoplasma gondii increases protein synthesis in infected macrophages.During infectious diseases, translational control can act as a host defense mechanism but also can be exploited by the invading pathogen as a survival strategy (30–32). To explore these possibilities during T. gondii infection, we inoculated bone marrow-derived murine macrophages (BMDM) with the RH (type I) and ME49 (type II) strains and assessed their effects on global protein synthesis by comparing the amounts of monosomes (inefficient translation) and heavy polysomes (efficient translation), as observed from polysome tracings. This approach revealed an increase in the amount of heavy polysomes with a concomitant decrease in the amount of monosomes in T. gondii-infected cells (Fig. 1A, left). The ratio of the normalized absorbance (254 nm) of heavy polysomes to monosomes indicated that type I and II strains enhance host cell protein synthesis to similar extents (mean fold changes over the value for the uninfected control [standard deviations {SD}] of 1.59 [0.19] for RH and 1.60 [0.11] for ME49) (Fig. 1A, right). These observed differences did not result from a contaminating signal originating from the parasite, as polysome tracings from T. gondii tachyzoites alone were barely detected (see Fig. S1A in the supplemental material). Thus, T. gondii infection leads to increased macrophage protein synthesis.

T. gondii infection stimulates protein synthesis and selectively modulates translational efficiencies in BMDM. (A) BMDM cultures were inoculated with either RH or ME49 T. gondii tachyzoites (MOI of 3:1) or left uninfected (control) for 8 h. Cell lysates were sedimented on 5 to 50% sucrose gradients. (Left) Gradients were fractionated, and the absorbance at 254 nm was recorded continuously. Absorbance values were normalized. Arrows indicate the 40S and 60S ribosomal subunits and 80S (monosomes). The light and heavy polysome regions were identified as fractions containing mRNA associated with 2 to 3 and >3 ribosomes, respectively. Polysome tracings are representative of data from at least four independent experiments. (Right) The area under the curve of the monosome and heavy polysome regions was calculated, and the heavy polysome-to-monosome ratios were then normalized to values for uninfected BMDM cultures (control) (means [SD]; n = 4 biological replicates). (B) Workflow strategy to characterize the translatome of T. gondii-infected BMDM. Cytosolic extracts from control (uninfected) and T. gondii-infected (RH or ME49) BMDM cultures were sedimented on a sucrose gradient. Heavy polysome fractions were pooled (referred to as heavy polysome-associated mRNA). Total and heavy polysome-associated mRNAs were sequenced by RNA-Seq and analyzed by ANOTA, an R package (n = 3 biological replicates). Validation of genes identified by RNA-Seq was accomplished with targeted nCounter assays (custom panel; n = 3 biological replicates). (C) Densities of FDRs for changes in polysome-associated mRNA, cytosolic mRNA, or translational efficiency (i.e., following ANOTA) for each strain compared to the uninfected control. (D) Scatter plots of log2 fold changes (for the same comparisons as in panel B) for polysome-associated and cytosolic mRNAs. Transcripts showing altered translational efficiencies and mRNAs showing a changed polysome association (which can be explained by altered cytosolic mRNA levels following, e.g., modulated transcription) are indicated. (C and D) Data analyses were performed on samples generated from three independent biological replicates.

T. gondii infection causes selective modulation of host cell translation.The increased amount of mRNA associated with heavy polysomes following parasite infection is consistent with parasite-induced mRNA-selective and/or global changes in host cell translational efficiency. To resolve the selective regulation of translational efficiency, we used polysome profiling, quantified by transcriptome sequencing (RNA-Seq) (Fig. 1B). During polysome profiling, both efficiently translated mRNA (defined here as mRNA associated with >3 ribosomes) and cytosolic mRNA (input) are isolated and quantified. As altered cytosolic mRNA levels will lead to corresponding changes in polysome-associated mRNA even without a modulation of translational efficiency, differences in polysome-associated mRNA levels require adjustment for changes in cytosolic mRNA levels to identify bona fide changes in translational efficiency. Here, we used the ANOTA (analysis of translational activity) algorithm for this purpose (45). Polysome profiling coupled with ANOTA of uninfected and RH- or ME49-infected BMDM revealed widespread selective alterations in translational efficiency upon infection with either strain (Fig. 1C). In total, 322 and 625 mRNAs showed activated and repressed translational efficiencies, respectively, in response to either strain (false discovery rate [FDR] of ≤0.25) (Fig. 1D; see also Table S1 in the supplemental material). In contrast, only 51 targets showed translational efficiencies modulated differently between strains RH and ME49 (Fig. S1B and Table S1). We selected 220 targets regulated via translation and 30 nonregulated genes for validation using a custom-designed nCounter code set from NanoString Technologies (Table S2). Targets were selected based on biological functions of the encoded proteins that were of particular interest (e.g., cell death, metabolism, the immune response, and translation). In addition, expression levels (similar numbers of low, intermediate, and high levels of polysome-associated mRNAs) and proportions of the directionality of translational regulation (∼1/3 up- and ∼2/3 downregulated) were also considered selection criteria. Changes in translational efficiency as estimated by RNA-Seq and nCounter assays were highly correlated (Spearman rank correlation, ρ, values of 0.76 for RH versus the control, 0.76 for ME49 versus the control, and 0.56 for RH versus ME49) (Fig. S1C). Accordingly, mRNAs identified as translationally activated or repressed using RNA-Seq largely showed a consistent directionality of regulation. Thus, both T. gondii strains induce abundant and similar selective changes in the host cell translational efficiency.

T. gondii selectively reprograms translation of mRNAs encoding proteins involved in protein synthesis, cell activation, and cytoskeleton/cytoplasm organization.To identify functional classes enriched for translationally regulated host mRNAs and to predict biological processes consequently affected during T. gondii infection, we used Ingenuity Pathway Analysis (IPA) (46). Within the molecular and cellular functions IPA classification, several functional networks were significantly enriched (FDR of <0.05) among proteins encoded by mRNAs whose translation was altered during T. gondii infection relative to the background (all mRNAs detected by RNA-Seq described above) (Fig. 2A; see also Table S3 in the supplemental material). The enriched categories include mRNAs involved in translation, the organization of the cytoskeleton and cytoplasm, cell activation, and cell movement. Globally, translational efficiencies of targets that belong to these categories displayed similar directionalities of regulation by RH and ME49 (Fig. 2B). Within the “translation” category, all mRNAs encoding ribosomal proteins were translationally activated. Conversely, targets grouped in the category “organization of the cytoplasm and cytoskeleton” were generally repressed. These transcripts encode proteins involved in membrane dynamics, microtubule-based movement, organelle formation and organization, and vesicular trafficking (e.g., dyneins Dync1H1 and Dync2H1; kinesins Kif11 and Kif3A; and myosins Myh10, Myo6, Myo7A, Myo10, and Myo18A). Within the category “activation of cells,” translational efficiencies of some mRNAs related to cell movement and chemotaxis were activated (chemokines Ccl5 and Ccl9), while others were repressed (chemokine receptors Ccr2, Ccr5, and CxcCr1 and matrix metalloproteases Mmp9 and Mmp13). Although not classified in this category by IPA, Mmp8, Mmp11, and Mmp12 were also identified as translationally downregulated upon infection (Table S1). Notably, mRNAs encoding immune cell activators were largely repressed (CD molecules Cd2, Cd93, and Cd180; interleukin receptors Il2Rb1, Il18Rap, and Il6Ra; and integrins Itgam and Itgav). In stark contrast, transcripts involved in immune suppression and negative regulation of inflammation (e.g., Cd200 and Il1Ra) were more efficiently translated in infected cells (Fig. 2B). Interestingly, the translational repression of transcripts promoting cell death was concomitant with the translational activation of antiapoptotic and cell cycle progression regulators. For instance, the levels of the negative regulator of cell differentiation Notch2 and the proapoptotic transcription factor Foxo3a were decreased, whereas the levels of antiapoptotic and positive regulators of cell cycle progression and macrophage differentiation were enhanced (e.g., Bcl2A1D and Hbegf) (Fig. 2B). Consistent with this, other negative regulators of cell proliferation and differentiation (Bcl6 and Notch4) were also translationally repressed (Table S1). Thus, T. gondii appears to selectively modulate the translational efficiencies of host mRNAs favoring the survival of infected cells while reorganizing the host cell architecture and preventing deleterious immune responses.

T. gondii infection selectively activates mTOR-sensitive translation. (A) Cumulative frequency distributions of RefSeq 5′ UTR lengths across the mm10 genome (black), compared to those of the upregulated and downregulated sets in RH (top) and ME49 (bottom) infections. (B) Sequence logo of the MEME motif found in the upregulated 5′ UTR set. Nucleotide positions are indicated on the x axis. (C) Venn diagrams indicating the numbers of genes harboring a TOP motif present in the upregulated versus downregulated gene sets. (D) Cumulative distribution of translational efficiencies of TOP mRNAs, IRES-containing mRNAs, and the background (all transcripts). (E) Cumulative distribution of translational efficiencies of mRNAs related to mitochondrial translation, the electron transport chain, mitochondrial respiratory chain complex I, mitochondrial respiratory chain complex IV, the proton-transporting ATP synthase complex, and the background (all transcripts). (F) Cumulative distribution of translational efficiencies of mRNAs whose translation was previously defined as depending on mTOR activity and the background (transcripts not belonging to the gene signature). For panels D to F, changes in translational efficiencies between RH-infected (left) or ME49-infected (right) BMDM and uninfected cells were evaluated. P values determined by a Wilcoxon test are indicated (tests consider genes belonging versus those not belonging to each group). ECDF, empirical cumulative distribution function. For panels A to F, data analyses were conducted on samples obtained from three independent biological replicates.

The data described above indicate that increased mTOR activity underlies a substantial proportion of parasite-induced changes in the translatome. To directly assess this, we used a previously reported translation signature (53), consisting of mRNAs whose translation efficiency is suppressed following mTOR inhibition (using PP242 in the breast carcinoma cell line MCF-7). Indeed, infection with either strain was associated with selectively activated translation of such mTOR-sensitive mRNAs (P < 0.001 for RH and ME49 versus the control) (Fig. 3F and Table S5). Altogether, these data suggest that T. gondii selectively augments mTOR-sensitive translation in BMDM.

T. gondii infection increases host mTORC1 activity in BMDM.The translation of mRNAs whose translational efficiency parallels mTOR activity was enhanced in T. gondii-infected macrophages. Therefore, we next determined mTORC1 kinase activity by assessing the phosphorylation of its downstream targets, S6K1/S6K2 and 4E-BP1/4E-BP2. Of note, parasite extracts (i.e., devoid of any host cell [“Tg only”]) were probed in parallel to rule out the possibility that the observed changes in mTORC1 signaling were due to cross-reactivity of the antibodies against parasite proteins (Fig. 4A). Phosphorylation of S6K1 (T389) and levels of S6K2 were increased in BMDM infected with either the RH or ME49 strain (Fig. 4A). Accordingly, the phosphorylation of the downstream target of S6K1/S6K2, ribosomal protein S6 (RPS6) (S235, S236, S240, and S244), was augmented and required active invasion without noticeable bystander effects (see Fig. S3A in the supplemental material). Phosphorylation of RPS6 in infected cells was completely abrogated in S6K1−/−/S6K2−/− BMDM, confirming that this cellular response is S6K1/S6K2 dependent (Fig. S3B). In addition, the hierarchical phosphorylation of 4E-BP1/4E-BP2 (i.e., T37/46, T70, and S65) was induced in T. gondii-infected BMDM with similar kinetics for the two strains (Fig. 4A). Hyperphosphorylation of 4E-BP1/4E-BP2 was also evident by a mobility shift on SDS-PAGE gels and increased amounts of the γ form compared to the α and β forms. The hyperphosphorylated forms of 4E-BP1/4E-BP2 have lower affinities for the mRNA 5′-m7G cap-binding protein eIF4E (29). Consistently, m7GTP pulldown assays revealed a reduced interaction between 4E-BP1/4E-BP2 and m7GTP-bound eIF4E in T. gondii-infected BMDM (Fig. 4B and C). Treatment with Torin-1, an active-site TOR inhibitor (asTORi) (54), resulted in the complete dephosphorylation of 4E-BP1/4E-BP2 and strong binding to m7GTP-bound eIF4E regardless of the infection status of the host cell (Fig. 4B). In summary, T. gondii activates mTORC1 signaling and promotes the dissociation of 4E-BP1/4E-BP2 from cap-bound eIF4E in BMDM.

T. gondii augments mTORC1 activity and promotes 4E-BP1/4E-BP2 dissociation from cap-bound eIF4E in BMDM. (A) BMDM cultures were inoculated with either RH or ME49 T. gondii tachyzoites or left uninfected for the indicated times. Phosphorylation and expression levels of indicated proteins were monitored by Western blotting. Total amounts of β-actin were used as a loading control, and an antibody against T. gondii profilin-like protein was employed to assess infection of BMDM cultures. Total protein extracts from extracellular tachyzoites (both the RH and ME49 strains) (Tg only) were used to control for any cross-reactivity of the antibodies against T. gondii proteins. (B) BMDM cultures were pretreated with 200 nM Torin-1 or an equal volume of DMSO (vehicle) for 2 h and then inoculated with either RH or ME49 T. gondii tachyzoites or left uninfected. Total protein extracts were collected at 8 h postinfection and prepared for m7GTP pulldown assays. Levels of the indicated proteins in the input (15 μg), pulled-down material (20%), and flowthrough (FT) (15 μg) were determined by Western blotting. (C) Ratios of m7GTP-bound 4E-BP/eIF4E were calculated based on densitometric measurements of the band intensities of 4E-BP1, 4E-BP2, and eIF4E and normalized to values for the uninfected BMDM control (means [SD]; n = 3 biological replicates). For panels A and B, data are representative of results from at least three independent biological replicates. **, P < 0.01.

Next, we aimed to elucidate upstream events activating mTORC1 during T. gondii infection. To this end, we assessed the contribution of AKT, which phosphorylates and inactivates the mTORC1 repressor TSC2 (55). Pretreatment with a pan-AKT inhibitor, MK-2206 (56), partially decreased the phosphorylation of RPS6 and 4E-BP1 (Fig. S3C). In light of these results, we monitored mTORC1 activity in T. gondii-infected BMDM deprived of l-leucine, an amino acid whose intracellular levels regulate mTORC1 in an AKT-independent fashion (57). Simultaneous pharmacological inhibition of AKT and l-leucine starvation abrogated RPS6 and 4E-BP1 phosphorylation in T. gondii-infected BMDM (Fig. S3C). Altogether, these data suggest that the activation of mTORC1 in T. gondii-infected BMDM relies on both AKT-dependent and -independent mechanisms.

Inhibition of mTOR reverses T. gondii-induced activation of host mRNA translation and dampens parasite replication.To further assess whether changes in translational efficiency observed in T. gondii-infected cells depend on mTOR, polysome tracings were generated from BMDM treated with rapamycin, an allosteric mTORC1 inhibitor, or Torin-1. Rapamycin induced similar shifts from heavy polysomes to monosomes in RH- and ME49-infected BMDM (48.3% and 37.2% reductions, respectively) (Fig. 5A and B). As expected, Torin-1 exerted a more dramatic effect than rapamycin (RH, 62.2%; ME49, 62.5%), owing to its greater potency (54). Of note, no differences in infection rates and no acute toxicity were observed for parasites exposed to the inhibitors (see Fig. S4 in the supplemental material). While rapamycin and Torin-1 abrogated the phosphorylation of S6K1 (T389) and its downstream target RPS6 (S235, S236, S240, and S244) in T. gondii-infected BMDM, only Torin-1 completely abolished 4E-BP1/4E-BP2 phosphorylation (as described previously [58]) (Fig. 5C). To evaluate whether chemical inhibition of mTOR is sufficient to reverse the translation program induced following parasite infection, we performed polysome profiling, quantified by nCounter assays (same targets as the ones described above) (Table S2), in T. gondii-infected and uninfected BMDM treated with rapamycin or Torin-1. We next assessed how fold changes in polysome-associated and cytosolic mRNA levels (for the subset of mRNAs translationally activated upon infection by either RH or ME49) were affected in cells treated with rapamycin or Torin-1. This approach revealed that chemical inhibition of mTOR is sufficient to largely reverse the effects on selective translation induced during parasite infection (Fig. 5D and Table S6).

Inhibition of mTOR reverses T. gondii-induced activation of host mRNA translation and dampens parasite replication. BMDM cultures were pretreated with 20 nM rapamycin, 200 nM Torin-1, or an equal volume of DMSO (vehicle) for 2 h and then inoculated with either RH or ME49 T. gondii tachyzoites or left uninfected (control) for 8 h. (A) Polysome tracings representative of data from at least four independent biological replicates. a.u., arbitrary units. (B) Heavy polysome-to-monosome ratios normalized to values for DMSO-treated uninfected BMDM cultures (control) (means [SD]; n = 4 biological replicates). (C) A fraction of the cultures utilized for polysome tracings was used to collect total protein extracts. The phosphorylation status and expression levels of the indicated proteins were monitored by Western blotting. Data are representative of results from four separate experiments. (D) Transcripts showing activated translational efficiencies following parasite infection were selected, and their mTOR-sensitive translation was assessed by using polysome profiling coupled to nCounter assays (same treatments as for panel A). The cumulative distributions of log2 fold changes for these targets were compared to the background (all other nCounter-quantified mRNAs) in infected cells with or without rapamycin or Torin-1 treatment and compared to values for uninfected DMSO controls. Empirical cumulative distribution functions (ECDF) are shown for log2 fold changes for cytosolic mRNA and polysome-associated mRNA. UP, upregulated. P values for comparison of selected targets with their corresponding background, determined by Wilcoxon tests, are given. Data analyses were conducted on samples obtained from three independent biological replicates. (E) BMDM were pretreated with 20 nM rapamycin, 200 nM Torin-1, or an equal volume of DMSO (vehicle) for 2 h and then inoculated with either RH or ME49 T. gondii tachyzoites at a parasite-to-BMDM ratio of 1:20 for 72 h. Genomic DNA was extracted, and parasite replication was assessed by qPCR by amplification of the T. gondii B1 gene. CT values were normalized to the value for mouse β-actin, and effects of the inhibitors on parasite replication were calculated by normalization to values obtained from cultures treated with DMSO for each strain. Values from one representative experiment of two independent trials are presented as means (SD); experiments on all samples were performed in technical triplicates. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

As parasite infection appears to profoundly affect host cell properties essential for the synthesis of biomolecules and growth by perverting host mTOR-sensitive mRNA translation, we hypothesized that the upregulation of mTOR activity constitutes a parasite survival strategy. To test this, BMDM were pretreated with either rapamycin or Torin-1 and infected with either RH or ME49 T. gondii tachyzoites. Parasites were allowed to egress and reinfect new host cells for 72 h, after which parasite numbers were evaluated by quantitative PCR (qPCR). Rapamycin treatment had a modest effect on parasite proliferation (12.8% and 19.5% reductions for RH and ME49, respectively), while Torin-1 caused a more pronounced reduction in replication (57.1% and 55.1%, respectively) than did dimethyl sulfoxide (DMSO) treatment (Fig. 5E). These data indicate that T. gondii relies on augmenting mTOR activity for unhindered replication.

DISCUSSION

Translational control of gene expression is a central mechanism of host defense (31, 32). As such, pathogens hijack the host translation machinery to survive (31, 32). Using transcriptome-wide polysome profiling, we demonstrate that the protozoan parasite T. gondii selectively augments mTOR-sensitive translation to favor host cell survival and parasite replication. Type I and II T. gondii strains regulate mTOR activity and mRNA translation in similar fashions, suggesting that this is a core process needed for parasite propagation that does not depend on strain-specific virulence factors. Further supporting the notion that regulation of mTOR-sensitive mRNA translation constitutes a parasite survival strategy, mTOR inactivation dampened T. gondii replication.

Our data on the activation of mTOR signaling are consistent with those of a recent analysis of the phosphoproteome of T. gondii-infected human foreskin fibroblasts (HFF) showing an enrichment of phosphorylated proteins both up- and downstream of mTOR, including RPS6 and 4E-BP1 (59). Interestingly, Wang and colleagues had previously reported that the phosphorylation of S6K1 following T. gondii infection was not consistently observed in different host cell lines (HFF, 3T3, and HeLa) (60–62). In some cases, no increased or prolonged phosphorylation of S6K1 was observed (60, 61), while in others, sustained and augmented phosphorylation was detected (62). To resolve the S6K-dependent phosphorylation of S6 in our experimental setting, we combined a chemical approach (i.e., mTOR inhibitors) and a genetic approach (i.e., BMDM derived from s6k1/s6k2 double-knockout [DKO] mice) that confirmed the requirement for S6K1 and S6K2 activity for RPS6 phosphorylation in T. gondii-infected macrophages. Our results showing inducible and sustained phosphorylation of 4E-BP1 following infection are also in sharp contrast to those of Wang and colleagues, who reported the absence of this phenomenon in T. gondii-infected 3T3 and HeLa cells (60, 61). This discrepancy might be related to distinct signals triggered by T. gondii depending on the species and/or the type of host cell, which could result in the differential activation of mTOR effectors. Further characterization of mTOR outputs in immune and nonimmune cell populations will shed light on this matter. The mechanism by which T. gondii upregulates mTOR activity appears to be multifactorial, as indicated by the requirement for both AKT-mediated signaling and AKT-independent events, such as l-leucine availability. Amino acid abundance promotes Rag-dependent translocation and activation of mTORC1 at the lysosomal surface (63). Notably, mTOR associates with the parasitophorous vacuole (PV) in T. gondii-infected cells (60). Those observations, along with our data, suggest that the PV surface offers a favorable site for mTORC1 activation during T. gondii infection.

Chemical inhibition of mTOR allowed us to confirm its requirement for selective translational activation in T. gondii-infected macrophages. Consistent with this, numerous mRNAs containing a 5′ TOP motif were present exclusively among translationally activated mRNAs, including those encoding ribosomal proteins. Interestingly, several of them play a role in cell cycle progression (e.g., RPS3, RPS7, RPS15a, RPL15, and RPL36a) and the regulation of apoptosis (e.g., RPS14, RPS29, and RPL7) (64). Moreover, T. gondii suppressed the translation of macrophage mRNAs encoding proapoptotic proteins, such as Foxo3a, while leading to the translational activation of prosurvival factors and cell cycle regulators (e.g., Bcl2A1D, Cdk2, Hbegf, and Id2). T. gondii has the ability to inhibit apoptosis and regulate the host cell cycle to promote infection (65–68). Moreover, mTOR activity is required for host cell cycle progression during T. gondii infection (60). In view of those studies and our present findings, T. gondii appears to regulate the translational efficiency of select mRNAs that favor the survival of the infected cell and ultimately enhance parasite replication.

Enrichment for transcripts encoding proteins related to cell activation and movement was mainly associated with translational repression by T. gondii. Strikingly, several of these factors are critical during toxoplasmosis. For instance, mice deficient in Ccr2 fail to recruit Gr1+ inflammatory monocytes to the intestine and are susceptible to oral challenge with T. gondii cysts (69). Similarly, genetic deletion of Ccr5 increases susceptibility to toxoplasmosis, and signaling through this receptor is required for interleukin-12 (IL-12) production by dendritic cells in response to T. gondii (70). In stark contrast, Cd200, whose translational efficiency was activated in T. gondii-infected macrophages, has a detrimental effect during chronic toxoplasmosis. Indeed, CD200 expression downregulates microglial activation in the brain of T. gondii-infected mice. Accordingly, Cd200−/− mice display an efficient local antiparasitic response (71). Those studies, along with our data, indicate that selective translational control of immune-related genes by T. gondii contributes to its ability to subvert protective immune responses.

Numerous mRNAs pertaining to mitochondrial functions were translationally activated upon T. gondii infection, including mitoribosomal proteins, factors implicated in the electron transport chain, and ATP synthases. These data are consistent with the mTOR-sensitive translation of mRNAs encoding proteins related to mitochondrial biogenesis and functions (72, 73). With regard to T. gondii, it is well documented that the PV associates with the host mitochondria (74–76). Accumulating evidence indicates that this association not only provides the parasite with nutrients, such as amino acids and glucose (77), but also leads to altered oxidative phosphorylation (OXPHOS) (78) and impacts innate immune responses (79). Moreover, the metabolism of intracellular T. gondii tachyzoites appears to rely on host-derived ATP (80), and parasite egress is particularly sensitive to host cell ATP levels (81). Therefore, it is plausible that the translational control of select host mRNAs by T. gondii contributes to the subversion of host mitochondrial functions, which in turn favors parasite metabolic activity and replication. Future studies should therefore aim to address these issues.

Several genes related to autophagy were identified in our study as translationally repressed in T. gondii-infected cells (e.g., Atp13a2, Htr2b, Iigp1, Kdr, Lrrk2, Nbr1, Tecpr1, Tpcn1, Trim21, and Wdfy3), while others were activated (e.g., Hspa8 and Mefv). Interestingly, host autophagy appears to have a dual role during T. gondii infection since it provides macromolecules to replicating parasites (82) but also contributes to parasite clearance by stripping the PV membrane (83). TRIM20, encoded by the MEFV gene, is induced by gamma interferon (IFN-γ) and is involved in “precision autophagy,” by specifically targeting the inflammasome components NLRP3, procaspase 1, and NLRP1 to limit excessive inflammation (84). Therefore, by favoring the translational efficiency of MEFV, it possible that the parasite aims at limiting inflammasome activation, a mechanism required for host resistance against T. gondii (85, 86). Translational repression of Iigp1 and Trim21 could also favor parasite persistence within the infected cell. IIGP1 (or Irga6) is targeted and inactivated by the parasite virulence factor kinase complex ROP5/ROP18/GRA7 (87) to prevent immune-related GTPase (IRG)-mediated destruction of the parasite (88, 89). Similarly, TRIM21 accumulates at the PV membrane following IFN-γ stimulation to restrict parasite replication and modulate inflammatory cytokine production in T. gondii-infected cells (90). Moreover, Trim21-deficient mice are more susceptible to toxoplasmosis and display higher parasite burdens and lower proinflammatory cytokine levels. Hence, it is tempting to suggest that modulation of translational efficiency of host mRNAs involved in autophagy constitutes an additional mechanism employed by T. gondii to promote infection.

In addition to the identification of pyrimidine-rich (CU) motifs in the 5′ UTR of mRNAs translationally activated upon T. gondii infection, 5′ UTR analysis indicated the presence of GC-rich stretches in ∼50% of the mRNAs that are translationally repressed. Of interest, this feature was absent in the mRNAs whose translation was activated upon parasite infection. GC-rich motifs in the 5′ UTR are associated with the formation of secondary structures that limit mRNA translation efficiency (91, 92). Translational control is also achieved via trans-regulatory factors, such as RNA-binding proteins, microRNAs, and “specialized ribosomes” (93–95). A recent study by Fischer and colleagues reported that infection by T. gondii causes the formation of stress granules containing poly(A)-binding proteins (PABPs) in the nucleus of the host cell (96). PABPs regulate several steps in mRNA processing, such as export, degradation, stability, polyadenylation, and deadenylation (97, 98). Upon stress (e.g., UV irradiation or viral infection), mRNA can be stored in nuclear stress granules through interactions with PABPs leading to translational silencing (99). Further investigation is needed to define the role of PABPs in translational control during T. gondii infection. It is conceivable that numerous mechanisms act in concert during T. gondii infection to fine-tune selective host mRNA translation.

Collectively, this study defines the translational signature of macrophages infected by T. gondii and provides evidence that translational control constitutes a key mechanism of host cell subversion by the parasite. Further supporting a central role for posttranscriptional regulatory mechanisms of gene expression during toxoplasmosis, a lack of correlation between transcriptomic and proteomic data was reported for human fibroblasts infected by T. gondii (100). Thus, it is likely that by modulating translation efficiencies in different cell types, T. gondii promotes its survival and dissemination through the infected host. Future studies using in vitro and in vivo models of T. gondii infection will contribute to a better understanding of the impact and the mechanisms underlying the regulation of mRNA translation during toxoplasmosis.

MATERIALS AND METHODS

Reagents.Culture media and supplements were purchased from Wisent and Gibco; cycloheximide (CHX) was purchased from BioShop; CellTracker green CMFDA (5-chloromethylfluorescein diacetate) dye was purchased from Molecular Probes; RNasin was provided by Promega; rapamycin (sirolimus) was bought from LC Laboratories; Torin-1 and MK-2206 were purchased from Cayman; pyrimethamine was obtained from Tocris; XTT [2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide] was ordered from Biotium; [5,6-3H]uracil was purchased from Perkin-Elmer; cOmplete EDTA-free protease inhibitor and PhosSTOP phosphatase inhibitor tablets were purchased from Roche; DAPI (4′,6-diamidino-2-phenylindole dilactate) and CellTracker green (CMFDA) were acquired from Invitrogen; and primary and secondary antibodies for Western blotting and microscopy were purchased from Cell Signaling Technologies, Santa Cruz Biotechnology, R&D Systems, Sigma-Aldrich, BioLegend, and Invitrogen.

Ethics statement.Experiments were carried out under a protocol approved by the Comité Institutionnel de Protection des Animaux of the INRS-Institut Armand Frappier (CIPA number 1502-03). This protocol respects procedures on good animal practice provided by the Canadian Council on Animal Care.

Infection of BMDM.Macrophages were plated 1 day before infection with T. gondii in BMDM culture medium without LCCM and allowed to adhere overnight at 37°C with 5% CO2. Cultures were serum starved for 2 h and treated with inhibitors, when applicable. BMDM were then inoculated with parasites (RH or ME49) or left uninfected in fresh medium with 1% FBS. Any remaining extracellular parasites were rinsed away with warm PBS (pH 7.2 to 7.4) 1 h following inoculation; fresh medium was added with inhibitors, when applicable; and cells were incubated until the end of the experiment. Unless specified otherwise, this procedure was carried out for all in vitro experiments.

Purification of RNA.Efficiently translated mRNA (associated with ≥3 ribosomes) and the input material (total cytoplasmic mRNA) were extracted with TRIzol (Invitrogen) and chloroform, followed by isopropanol precipitation. In brief, polysome fractions and input material were diluted up to 1 ml with RNase-free H2O, and 500 μl of TRIzol and 150 μl of chloroform were then added. Samples were vortexed vigorously for 5 s, incubated for 3 min at RT, and centrifuged (18,000 × g for 15 min at 4°C). The aqueous phase of each sample was mixed with an equal volume (∼750 μl) of ice-cold isopropanol and 1 μl of GlycoBlue (Thermo Fisher), and the RNA was then precipitated overnight at −20°C and pelleted by centrifugation (18,000 × g for 10 min at 4°C). The RNA pellet was washed twice with 500 μl of 75% ice-cold ethanol. Samples were air dried and resuspended in RNase-free H2O, and polysome fractions from the corresponding treatment were pooled. RNA was further cleaned by using an RNeasy MinElute cleanup kit (Qiagen), according to the manufacturer's specifications. Finally, the approximate concentration, purity, and integrity of the purified RNA were assessed spectrophotometrically by using a NanoDrop 1000 instrument (Thermo Fisher) and a Bioanalyzer 2100 instrument with a eukaryote total RNA nanochip (Agilent Technologies). Samples were aliquoted and stored at −80°C until further analyses.

RNA-Seq data processing.Library preparation and sequencing were carried out at SciLifeLab (Stockholm, Sweden). RNA-Seq libraries were generated by using the strand-specific TruSeq protocol and sequenced by using an Illumina HiSeq2500 instrument with a single-end 51-base sequencing setup for both total cytosolic and polysome-associated mRNAs (mRNAs associated with >3 ribosomes) from three independent biological replicates. RNA-Seq reads from infected samples were first aligned to T. gondii reference genomes of GT1 (type I) and ME49 (type II) (the genome of the RH strain is not completely annotated, but it was estimated that among T. gondii RH gene families, 97.8% have orthologs in the GT1 strain [102]). Reads from infected samples that did not map to the T. gondii genomes as well as all reads from uninfected samples were aligned to the mouse genome mm10. HISAT2 was used for all alignments, with default settings (103). Alignments to T. gondii and Mus musculus resulted in 4.4% and 93.7% mappings on average, respectively. Gene expression was quantified by using the RPKMforgenes.py script (http://sandberg.cmb.ki.se/media/data/rnaseq/rpkmforgenes.py) (104) with options -fulltranscript -readcount -onlycoding, from which raw per-gene RNA-Seq counts were obtained (version last modified 2 July 2014). Annotation of genes was done by using RefSeq. Genes that had zero counts in all samples were removed. One replicate from the uninfected control condition was excluded from downstream analysis based on outlier behavior in principal-component analyses.

RNA-Seq analysis of the empirical data set using ANOTA.The limma::voom R function was used to compute log2 counts per million (105). To identify changes in translational efficiency leading to altered protein levels, ANOTA (45, 106) was used with the following parameters: minimum slope (minSlope) of −0.5 and maximum slope (maxSlope) of 1.5. Replicate was included as a batch effect in the models. The following criteria were considered for significant changes in translational efficiency: a false discovery rate (FDR) of <0.25, a translational efficiency (apvEff) of >log2(1.5), a log ratio of polysome-associated mRNA data to cytosolic mRNA data (deltaPT) of >log2(1.25), and a polysome-associated mRNA log fold change (deltaP) of >log2(1.5). Genes that overlap, resulting in RNA sequencing reads that cannot be associated with only one gene, were excluded from downstream analyses. Similar FDR and fold change thresholds were used when selecting differential expression of polysome-associated mRNAs. For further analysis, we used a previously reported list of TOP- and IRES-containing mRNAs (107), mitochondrion-related function genes included in the Gene Ontology (GO) (2015) terms indicated in Table S5 in the supplemental material, and a signature for mTOR-sensitive translation (53); gene symbols were converted from human to mouse and are listed in Table S5. The difference in log2 fold changes of translational efficiencies between the signatures and the background was assessed by using Wilcoxon-Mann-Whitney tests.

nCounter analysis.For nCounter analysis (NanoString Technologies), a subset of 250 genes was selected according to RNA-Seq data. Sequences for the probes in the custom-designed nCounter code set are listed in Table S2 in the supplemental material. Infected and uninfected cells were treated with rapamycin or Torin-1, followed by the isolation of efficiently translated mRNA and cytosolic mRNA from three independent biological replicates (as described above). The positive spike-in RNA hybridization controls for each lane were used to normalize NanoString counts as implemented in the posCtrlNorm function of the Bioconductor package NanoStringQCPro (108) (with parameter summaryFunction = “sum”). Targets with fewer than half of the samples above the background level (7 [log2 scale]) were excluded. The quality of the normalization was assessed by using 30 nonregulated genes selected from the RNA-Seq data analysis to verify that their expression levels remained stable in the DMSO samples. Spearman rank correlation coefficients were calculated between fold changes obtained from the RNA-Seq data and those obtained from the nCounter data. Analysis of translational control was performed with ANOTA to select targets that were translationally upregulated upon infection with each T. gondii strain. Genes with an FDR of <0.25 were considered significant, and the following parameters were used: minSlope of −1, maxSlope of 2, and selDeltaPT of log2(1.2). For these targets, empirical cumulative distribution function (ECDF) curves were used to visualize the distribution of their log2 fold changes (for both cytosolic and polysome-associated mRNAs) upon treatment with rapamycin or Torin-1. Distribution differences were evaluated by using Wilcoxon-Mann-Whitney tests.

Functional classification of gene lists.Enrichment of translationally regulated genes in specific functional networks was determined by using Ingenuity Pathway Analysis (IPA; Qiagen) by comparing ANOTA-regulated gene sets against the entire sequenced data sets (46). Within the IPA application, statistical significance was calculated by using a right-tailed Fisher exact test, and P values were adjusted for multiple-hypothesis testing by using the Benjamini-Hochberg method to arrive at a FDR.

RNA sequence motif analyses.For 5′ UTR sequence analysis, nonredundant RefSeq 5′ UTRs were retrieved from genome build mm10 using the UCSC table browser (https://genome.ucsc.edu/cgi-bin/hgTables). The length and GC content of the UTRs were computed by using a custom script. To minimize the bias associated with analyzing GC contents and sequence motifs of very short sequences, 5′ UTRs of <20 nucleotides were omitted from further analyses. Novel RNA sequence motifs were detected in the 5′ UTRs by using MEME (Multiple Em for Motif Elicitation) (48). To accurately estimate the probability of a candidate motif appearing by chance (E value) and to improve the sensitivity of the motif search, a custom background model was employed. Briefly, the “fasta-get-markov” utility included in the program was used to compute a background Markov model of order 1 using all RefSeq mouse 5′ UTRs. Sequence logos were generated by using the ggseqLogo package in R. Cap analysis of gene expression (CAGE)-Seq data were retrieved from FANTOM5 (http://fantom.gsc.riken.jp/5/), and sequences were mapped to the mm9 build by using the UCSC genome browser.

XTT and [5,6-3H]uracil incorporation viability assays.To test for acute toxicity of the inhibitors against extracellular tachyzoites, parasite viability was measured by using the tetrazolium reduction-based XTT assay (109). Briefly, freshly egressed tachyzoites (RH and ME49 strains) were resuspended in culture medium (without FBS) and transferred to 96-well plates (2 × 106 tachyzoites per well). Parasites were treated with 20 nM rapamycin, 200 nM Torin-1, or an equal volume of the vehicle (DMSO) and incubated for 2 h at 37°C with 5% CO2. As a negative control, heat-killed (HK) parasites incubated at 56°C for 10 min were included in the assay. Next, the XTT reagents were prepared according to the manufacturer's specifications (Biotium) and added to the parasites. Tachyzoite cultures were incubated for 20 h. Absorbance was measured by using a Multiskan GO 1510 plate reader (Thermo Fisher) at 470 nm, from which the absorbance at 660 nm was subtracted.

To test for acute toxicity of the inhibitors on intracellular tachyzoites, the incorporation of 5,6-3H-radiolabeled uracil by the parasite was measured as an assessment of viability (110). Briefly, BMDM were plated in 24-well plates overnight; serum starved for 2 h; treated with either 20 nM rapamycin, 200 nM Torin-1, or an equal volume of the vehicle (DMSO); and inoculated with T. gondii tachyzoites (RH and ME49 strains) at an MOI of 3:1 in fresh medium with 1% heat-inactivated dialyzed FBS. Cultures were incubated overnight, 5 μCi of [5,6-3H]uracil (Perkin-Elmer) was then added to each well, and cultures were incubated for another 2 h at 37°C with 5% CO2. Plates were chilled at −20°C for 2 min, an equal volume of ice-cold 0.6 N trichloroacetic acid (TCA) was added to each well, and cells were fixed on ice for 1 h. Wells were rinsed with PBS and then with deionized H2O. Plates were air dried, and the precipitated material was resolubilized in 500 μl of 0.1 N NaOH for 1 h. Half of the material was thoroughly mixed with an equal volume of OptiPhase Supermix scintillation cocktail (Perkin-Elmer), and radioactivity was measured (counts per minute) with a MicroBeta TriLux scintillation counter (Perkin-Elmer). Uninfected cells were included to measure the background incorporation of uracil by host cells, and treatment with 10 μM pyrimethamine was included as a positive control for chemical toxicity against T. gondii parasites.

Immunofluorescence microscopy.BMDM were seeded onto glass coverslips in 24-well plates overnight, serum starved for 2 h, and then inoculated with T. gondii tachyzoites (RH and ME49 strains), previously stained with 20 μM CellTracker green CMFDA dye (Molecular Probes) for 20 min, at an MOI of 3:1 or left uninfected. After 8 h of infection, cells were rinsed with PBS three times and then fixed with 3.7% paraformaldehyde (PFA) (in PBS) for 15 min at RT. Cell membranes were permeabilized with 0.2% Triton X-100 (in PBS) for 5 min at RT. Fc receptors and nonspecific binding sites were blocked by incubating samples with 5 μg/ml anti-CD16/32 (BioLegend) diluted in PBS supplemented with 10 mg/ml bovine serum albumin (BSA). Primary antibody against phospho-RPS6 (S240/244) (Cell Signaling Technologies) was diluted in PBS with 10 mg/ml BSA and incubated with samples for 1 h at RT. A goat anti-rabbit IgG(H+L) secondary antibody conjugated to Alexa Fluor 594 (Invitrogen) was incubated with the samples for 1 h at RT. Nuclei were stained with 300 nM DAPI (Invitrogen) for 5 min at RT. Coverslips were mounted onto slides with Fluoromount G (Southern Biotech). Samples were visualized by using a Leica microscope, and image processing was performed with Fiji (111).

Measurement of parasite replication by qPCR.Parasite replication in vitro was assessed by extracting genomic DNA (gDNA), followed by measuring the amplification of the T. gondii B1 gene by qPCR, as described previously (101). Briefly, BMDM cultures were plated in 12-well plates overnight; serum starved for 2 h in the presence of DMSO (vehicle), 20 nM rapamycin, or 200 nM Torin-1; and then inoculated with T. gondii tachyzoites (RH and ME49 strains) at an MOI of 1:20. Parasites were allowed to replicate, egress, and reinfect cells for 72 h. Culture supernatants were collected (for extracellular parasites), while the remaining adherent cells (containing intracellular parasites) were detached by adding ice-cold PBS with 5 mM EDTA for 15 min at 4°C, followed by vigorous pipetting, and then pooled into culture supernatants. Tachyzoites and host cells were pelleted by centrifugation (16,000 × g for 10 min at 4°C) and then resuspended in 200 μl PBS. gDNA was extracted by using a Roche High Pure PCR template preparation kit, according to the manufacturer's specifications. To measure parasite replication, the 35-fold repetitive T. gondii B1 gene was amplified by qPCR using the PowerUP SYBR green PCR master mix (Applied Biosystems) with the MgCl2 concentration adjusted to 3.5 μM, 10 ng of template gDNA, and 0.5 μM forward primer (5′-TCCCCTCTGCTGGCGAAAAGT-3′) and reverse primer (5′-AGCGTTCGTGGTCAACTATCGATTG-3′) (Integrated DNA Technologies) in a 20-μl reaction mixture volume. The reaction was carried out in a QuantStudio 3 real-time PCR system (Applied Biosciences) with the following program: 10 min of initial denaturation at 95°C, followed by 45 cycles of 15 s of denaturation at 95°C, 30 s of annealing at 58°C, and 30 s of extension at 72°C. Cycle threshold (CT) values were normalized by using mouse β-actin gene primers (0.2 μM forward primer 5′-CACCCACACTGTGCCCATCTACGA-3′ and reverse primer 5′-CAGCGGAACCGCTCATTGCCAATGG-3′), and the MgCl2 concentration was adjusted to 2.5 μM. Analysis was carried out by relative quantification using the comparative CT (2−ΔΔCT) method (112). The effect of the inhibitors on replication was calculated by normalization to values obtained from parasite cultures treated with DMSO. Values are presented as means (SD); experiments on all samples were performed in triplicates.

Statistical analysis.Where applicable, data are presented as means (SD). Statistical significance was determined by using one-way analysis of variance (ANOVA) followed by a Tukey post hoc test; calculations were performed by using Prism software (GraphPad, La Jolla, CA). Differences were considered significant when P values were <0.05, <0.01, or <0.001.

ACKNOWLEDGMENTS

We are grateful to Nahum Sonenberg for providing bone marrow from C57BL/6 s6k1−/− s6k2−/− mice (McGill University, Montreal, QC, Canada). We thank Medhi Jafarnejad (McGill University) for technical advice. We are thankful to Annie Sylvestre and Annik Lafrance for invaluable technical assistance. We acknowledge support from the Science for Life Laboratory, the National Genomics Infrastructure (NGI), and Uppmax for providing assistance in massive parallel sequencing and computational infrastructure.

This work was supported by a Basil O'Connor starter scholar research award (5-FY14-78) and a research grant (6-FY16-151) from The March of Dimes Foundation to M.J. The Centre for Host-Parasite Interactions is supported by a Subvention de Regroupement Stratégique from the Fonds de Recherche du Québec en Nature et Technologies (FRQ-NT). M.J. is a recipient of a Bourse de Chercheur-Boursier Junior 1 from the Fonds de Recherche du Québec en Santé (FRQ-S) and a Subvention d'Établissement de Jeune Chercheur from the FRQ-S. V.C. is supported by a Ph.D. scholarship from the Fondation Universitaire Armand Frappier. Research in the laboratory of O.L. is supported by grants from the Swedish Research Council and the Wallenberg Academy Fellows program. The funders had no role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript.

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